The human hepatoma cell line HepG2 were obtained from the American Type Culture Collection (ATCC; Manassas, VA, USA). The HepG2/HBx and HepG2/pcDNA3.1 cell lines, were derived from HepG2 cells by transfecting with HBx expression plasmid or an empty plasmid (pcDNA3.1(+)/V5-HisB), respectively. Both cell lines have been successfully established (27). All cell lines were cultured in DMEM (Gibco, Carlsbad, CA, USA) supplemented with 10% fetal bovine serum (FBS; Gibco, Grand Island, NY, USA) and maintained in humidified incubator at 37°C in a 5% CO₂ atmosphere.

The γ-secretase inhibitor N-[N-(3,5-difluorophenacetyl)-L-alanyl]-S-phenylglycine t-butyl ester (DAPT) was purchased from the Sigma-Aldrich Company (St. Louis, MO, USA). DAPT was dissolved in 100% dimethylsulphoxide (DMSO, Sigma) to make a stock solution of 10 mM, which was then diluted in culture medium to obtain the desired concentrations of 1, 5, 10 and 20 μM. DMSO diluted in culture medium at the final concentration of 0.05% without DAPT was designated as 0 μM. Untreated cells were those incubated in the culture medium without any additives. Cells treated with or without DAPT were cultured for 48 h, after which the total RNA or protein was extracted and flow cytometry was carried out.

Cell proliferation assays were performed by using a Cell Counting Kit-8 (Dojindo, Kumamoto, Japan) according to the manufacturer’s instructions. Briefly, 1×10⁴ cells/well were plated in 96-well plates and cultured with growth medium. At the indicated time points, the medium was aspirated. Then 100 μl serum-free DMEM and 10 μl WST-8 [2-(2-methoxy-4-nitrophenyl)-3-(4-nitrophenyl)-5-(2,4-disulfophenyl)-2H tetrazolium, monosodium salt] were added to each well followed by incubation at 37°C for 1.5 h. Absorbance was measured at 450 nm with a reference wavelength of 630 nm on a spectrophotometer (Molecular Devices, Sunnyvale, CA). To evaluate the viability of HepG2/HBx cells, 1×10⁴ cells/well were plated in 96-well plates followed 12 h later by the addition of DAPT at concentrations of 0, 1, 5, 10 or 20 μM. The cell viability was assessed as the percent of viable cells relative to the untreated control cells, which was determined for each concentration using the following equation: % viability = OD_experiment/OD_control × 100%. Control cells were considered as 100% viable. All experiments were repeated five times.

After treatment with or without DAPT for 48 h, cells were harvested, immediately fixed in 75% ethanol at 4°C overnight, treated with 50 mg/l RNAse A (Sigma) for 30 min at 37°C, and stained with 50 mg/l PI (Sigma) for 10 min. Samples were then analyzed for their DNA content by a FACSAria Cell Cytometer (BD Biosciences, San Jose, CA, USA). The data were analyzed with the CellQuest software (BD Biosciences). Apoptosis analysis was performed by using a Annexin-V-FITC kit (Bender MedSystems, Burlingame, CA, USA) according to the manufacturer’s instructions. The percentage of cells that were Annexin-V positive but PI negative was compared among the different treatment groups.

Immunofluorescence assays were performed as previously described (28). HepG2/HBx cells were cultured on glass coverslips for 24 h and fixed with 4% paraformaldehyde. The fixed cells were incubated with anti-HBx (1:200, Santa Cruz Biotechnology, Santa Cruz, CA) and anti-NICD (1:200, Santa Cruz Biotechnology) for 12 h at 4°C, then incubated with Cy3-conjugated goat anti-mouse IgG (1:100, Boster, China) and FITC-conjugated goat anti-rabbit IgG (1:100, Boster) for 1 h. Nuclei were stained with diamidinophenyl indole (DAPI) (Boster). The stained cells were observed using a Fluoview FV1000 laser scanning confocal microscope (Olympus, Japan).

Co-immunoprecipitation assays were performed as previously described (28). HepG2/HBx cells were lysed, and the lysates were pretreated with protein G-agarose (Santa Cruz Biotechnology) to remove non-specifically bound proteins. After centrifugation, one third of the supernatants were stored at −80°C as a positive control. The remaining supernatant samples were incubated for 2 h at 4°C with 1 μg of non-immune mouse IgG or mouse anti-HBx (Santa Cruz Biotechnology), for the negative control and the experimental group, respectively. Then, the mixture was incubated for 1 h to overnight at 4°C with 20 μl protein G-agarose (Santa Cruz Biotechnology). The immunocomplexes were extensively washed with PBS, samples were boiled in electrophoresis sample buffer and assayed by Western blot analysis.

Total RNA was isolated from cultured cells using TRIzol reagent (Invitrogen, Carlsbad, CA, USA) and cDNA was synthesized from 2 μg of total RNA using Moloney murine leukemia virus reverse transcriptase (MMLV) (Promega, Madison, WI, USA). Real-time quantitative PCR (qRT-PCR) using SYBR Premix (DRR041A, Takara, Japan) was performed as previously described (29). Amplifications were performed in a LightCycler machine (Roche Diagnostics, Basel, Switzerland) following the manufacturer’s instructions. The HepG2 cDNA was used as standard cDNA. A standard curve for each gene was generated from serially diluted standards, and values for unknown samples were extrapolated. β-actin was used as an internal control to normalize samples. All standards and samples were assayed in triplicate. The primer sequences used to amplify specific target genes are listed in Table I.

Cells were lysed as previously described (29) and the lysates were subjected to electrophoresis on SDS-PAGE and transferred to PVDF membranes (Millipore, Billerica, MA). The blotted membranes were blocked and subsequently incubated with rabbit anti-Jagged-1 (1:600), rabbit anti-Notch-1 (1:1,000), rabbit anti-Notch-1-IC (1:1,000), rabbit anti-Hes-1 (1:600), and rabbit anti-actin (1:1,000). All antibodies were from Santa Cruz Biotechnology except for anti-Notch-1-IC which was obtained from Cell Signaling Technology (Danvers, MA). After incubation with horseradish peroxidase-labeled secondary antibody (1:5,000–10,000; Santa Cruz Biotechnology), visualization was performed by an enhanced chemiluminescence kit (Pierce, Rockford, IL) and exposure to X-ray film (Kodak, Rochester, NY). Immunoblotting with the anti-actin antibody was used as an internal control to confirm equivalent protein loading. Each experiment was performed at least 3 times. The relative intensity of each protein band was scanned by Quantity One software (Bio-Rad Laboratories, Hercules, CA, USA).

SPSS version 17.0 software (SPSS for Windows, Inc., Chicago, IL, USA) was used for all statistical analyses. All results are expressed as mean ± SEM. Statistical analysis of the data was performed using a standard one-way ANOVA or one-way ANOVA for repeated measures, followed by the least significant difference (LSD) post-hoc test. Bonferroni’s correction was used to adjust for multiple comparisons. A 2-tailed Student’s paired t-test was also used to compare the differences in the values between two groups. A P-value <0.05 was considered to be statistically significant.

Previously, we observed that the HBx protein stimulated the proliferation as well as the cell cycle, inhibited the apoptosis of HepG2 cells significantly, and promoted tumour growth in nude mice (27). To investigate the potential involvement of Notch signaling in the development of HCC, we performed western blot analyses in HepG2, HepG2/pcDNA3.1 and HepG2/HBx cells. As shown in Fig. 1A, the protein levels of Jagged-1, Notch-1, Notch-1C and Hes-1 in HepG2/HBx cells were elevated relative to the controls. Then, we analyzed the mRNA levels of Jagged-1, Notch-1 and Hes-1 using quantitative real-time RT-PCR (qRT-PCR) and found that their mRNA levels were significantly increased in HepG2/HBx cells as compared with the controls (Fig. 1B).

To study the possible relationship between Notch-1 and HBx, we performed immunofluorescence and co-immunoprecipitation assays. As shown in Fig. 1C, the nuclei were stained blue (i), HBx was stained red (ii) whereas the NICD was stained green (iii) in HepG2/HBx cells. Yellow staining in the dual-labeling experiments indicates overlapping areas of red and green fluorescent labels (iv), suggesting co-localization of NICD with HBx in the HepG2/HBx cells. The compounds immunoprecipitated with anti-HBx or non-immune mouse IgG from HepG2/HBx cells were subjected to western blot analysis with anti-NICD and anti-Jagged-1, respectively. As shown in Fig. 1D, NICD was co-immunoprecipitated with HBx. No specific interaction was found between Jagged-1 and HBx or the protein immunoprecipitated by non-immune IgG, indicating the specificity of the NICD-HBx interaction.

In order to further establish that active Notch signaling is important for HBx to function as an oncoprotein, we treated HepG2/HBx cells with various concentrations of DAPT for 12 to 96 h and assessed cell proliferation by the WST-8 assay. As shown in Fig. 2A, in HepG2/HBx cells, increasing concentrations and durations of treatment with DAPT resulted in a decrease of cell viability. In contrast, treatment with various concentrations of DAPT for 12 or 24 h did not produce any significant reduction in cell viability. However, treatment of DAPT for >48 h resulted in significant dose- and time-dependent reduction in cell viability of HepG2/HBx cells. A significant reduction in cell viability by DAPT treatment was observed at concentrations of 10 and 20 μM after 48 h, with inhibition rates of 7.30 and 7.73%, respectively (P<0.05, Fig. 2A). We, therefore, selected the DAPT treatment time point of 48 h for further studies.

HepG2/HBx cells treated with various concentrations of DAPT for 48 h and assessed for the DAPT inhibitory effects on Notch-1 signaling by western blot analysis of Jagged-1, Notch-1, Notch-1-IC, Hes-1 protein levels, and by qRT-PCR of Hes-1 transcripts. As shown in Fig. 2B and C, DAPT treatment greatly reduced the amount of Notch-1-IC and Hes-1 protein in a dose-dependent manner, while there was no significant effect on Jagged-1 and Notch-1 levels (Fig. 2B). A significant reduction of Notch-1-IC and Hes-1 protein levels were observed at 10 and 20 μM of DAPT treatment. qRT-PCR experiments showed that the downregulation of Hes-1 transcripts in HepG2/HBx cells occurred after treatment with DAPT at 10 (P<0.05) and 20 μM (P<0.05) (Fig. 2C). These data were consistent with the previous results that treatment with 10 and 20 μM DAPT for 48 h significantly inhibited the proliferation of HepG2/HBx cells.

We performed PI staining and flow cytometry to define the cell-cycle distribution of DAPT-treated HepG2/HBx cells. Treatment of HepG2/HBx cells with 0–20 M of DAPT for 48 h resulted in arrest in the G0/G1 phase and shortening of the S phase. The proportion of HepG2/HBx cells in the G0/G1 phase was significantly increased at concentrations of 10 (88.99%, P<0.05) and 20 μM (90.95%, P<0.05), compared to that of 80.23% in untreated control cells (Fig. 3A and B). The proportion of cells in S phase was significantly decreased at concentrations of 10 (4.60%, P<0.05) and 20 μM (3.93%, P<0.05), relative to that of 12.55% in untreated control cells (Fig. 3A and B). No significant difference was observed between the G2/M phases. As shown in Fig. 3C and D, the apoptotic rates were increased from 5.45% in non-DAPT-treated cells to 13.28% at a concentration of 10 μM (P<0.01, Fig. 3C and D) and to (20.22%) at 20 μM (P<0.001, Fig. 3C and D) after treatment with DAPT for 48 h. Taken together, these results indicate that the inhibition of Notch signaling was associated with decreased DNA synthesis (S phase) and increased apoptosis, which contributed to the impaired growth of HepG2/HBx cells treated with DAPT.

As a control, Notch signaling was blocked in HepG2 cells using DAPT as well. HepG2 cells were treated with DAPT at the final concentration of 10 μM or with 0.05% DMSO. The proliferation rate of HepG2 cells was measured after DAPT treatment using WST-8 assay. No effect on cell proliferation could be observed in HepG2 cells upon ablation of Notch signaling (Fig. 4A). The protein expression level of Hes-1 was monitored using western blot analysis. As shown in Fig. 4B, treatment of HepG2 cells with DAPT led to a distinct Hes-1 downregulation after 48 h. These results demonstrated that Notch signaling was much less important for the growth of HepG2 cells than for that of HepG2/HBx cells.